Article pubs.acs.org/ac
Analysis of Soluble Lignin in Sugarcane by Ultrahigh Performance Liquid Chromatography−Tandem Mass Spectrometry with a Do-ItYourself Oligomer Database Eduardo Kiyota, Paulo Mazzafera, and Alexandra C. H. F. Sawaya* Plant Biology Department, Biology Institute, State University of Campinas, Unicamp, Campinas, São Paulo, 13083-970, Brazil S Supporting Information *
ABSTRACT: Lignin is a polymer found in the cell wall of plants and is one of the main obstacles to the implementation of secondgeneration ethanol production because it confers the recalcitrance of the lignocellulosic material. The recalcitrance of biomass is affected by the amount of lignin, by its monomer composition, and the way the monomers are arranged in the plant cell wall. Analysis of lignin structure demands mass spectrometry analysis, and identification of oligomers is usually based on libraries produced by laborious protocols. A robust method to build a doit-yourself lignin oligomer library was tested. This library can be built using commercially available enzymes, standards, and reagents and is relatively easy to accomplish. An ultrahigh performance liquid chromatography−tandem mass spectrometry (UPLC−MS/MS) method for the separation and characterization of monomers and oligomers was developed and was equally applicable to the synthetic lignin and to soluble lignin extracted from a sample of sugar cane.
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here is great interest in biomass as a source of storable liquid transportation fuels, thereby reducing the greenhouse gas emissions of fossil fuels.1,2 Potentially, ethanol can be produced from many plant sources. In Brazil, ethanol production is based on sucrose from sugar cane and in the United States on corn starch. Unfortunately, these are also important food sources, so fuel production may compete with food production. Alternatively, ethanol could be produced from the lignocellulosic biomass left over from commercial crops,3−5 such as sugar cane bagasse. Ethanol produced from lignocellulosic biomass is named second-generation ethanol. One of the main obstacles to the implementation of secondgeneration ethanol production is the recalcitrance of the lignocellulosic material. Bagasse, for example, is mainly composed of cellulose, hemicellulose, and lignin, which cannot be easily separated into usable components. Initially this material must be pretreated for lignin extraction to make cellulose accessible to further treatment.1,6 Cellulose is then hydrolyzed to release monomeric glucose, which in turn is fermented and converted into ethanol.7 Recalcitrance of the plant biomass is mainly due to the presence of lignin, a complex phenolic biopolymer, which is difficult to break down. The lignin content of biomass also limits its use as animal feed. Lignin biosynthesis is based mainly on the radical coupling of three hydroxyphenylpropanoid monomers (or monolignols), p-hydroxyphenyl (H), guayacyl (G), and syringyl (S), derived from phenylpropanoid alcohols (Figure 1), through a series of oxidations mediated by extracellular enzymes: peroxidase and laccase.8,9 Their individual contribution to lignin composition varies significantly © 2012 American Chemical Society
Figure 1. Structures of the three main lignin monomers: p-coumaryl alcohol (H), coniferyl alcohol (G), and sinapyl alcohol (S).
among species, cell types, and between tissues in the same plant.10 In addition to the three canonical monolignols, other phenylpropanoids can be incorporated into the polymer at varying levels, including hydroxycinnamates, hydroxycinnamyl aldehydes, and hydroxycinnamyl acetates.11 Plant biomass may be delignified by pretreatment with organic solvents, ionic liquids, Kraft pulping, dilute acid, ammonia fiber expansion, and hydrothermolysis,2 all costly methods that sometimes require high temperatures. While improvements in the pretreatments used to produce biofuels are being developed,12,13 genetic engineering of plants has been another approach, aiming to increase the efficiency of secondgeneration ethanol production by altering lignin composition and content in plants.14 Received: April 27, 2012 Accepted: July 24, 2012 Published: July 24, 2012 7015
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into the library, permitting their detection in plant varieties with greater digestibility, and which may consequently be economically more adequate for biofuel production as well as for animal feed. The fast chromatographic separation promoted by UPLC coupled to the richness of the MS/MS data permits the routine analysis of large numbers of samples. In the present study, soluble lignin oligomers from sugar cane were analyzed according to this procedure, and their composition and structure were determined by comparison to our do-it-yourself (DIY) library.
Lignin is involved in mechanical support, resistance to stress, pests, and disease,15 as well as water and nutrient transport;16 therefore, it cannot simply be eliminated (or even drastically reduced) in plants. Experiments down- and/or up-regulating specific genes involved in lignin biosynthesis and their impact on plant structure have been reported.2 Manipulation of key genes in tobacco plants,17−19 alfalfa, and grasses20−24 resulted in altered lignin composition and increased digestibility. Different phenylpropanoids can also be incorporated into the lignin structure with a marked increase of extraction efficiency.25 Another possible modification is related to the intermonomer linkages. The 8−O−4 linkage is the most common and more easily cleaved26 than other linkages, which are more chemically recalcitrant. It is believed that the relative proportion of monomers dictates the relative abundance of the interunit linkage present. For example, lignin rich in G units contains more recalcitrant 8−5, 5−5, and 5−O−4 links8 and lignin enriched with S units is less cross-linked and less recalcitrant to extraction. For this reason, lignin composition is classically described by the relative abundance and ratio of H, G, and S units.27 Lignin content is often measured by conventional gravimetric methods, with the most common being Klason and acid detergent fiber (based on the isolation of cell wall material), and spectrophotometric methods such as acetyl bromide method (AB) and thioglycolic acid method (TGA), which degrade the cell wall and lignin and determine soluble degradation products.28,29 However, different methods used for the determination of lignin in the same sample have resulted in different estimates of lignin concentration.29 Furthermore, these methods provide an estimate of the content and sometimes the composition of the insoluble lignin incorporated in the cell wall but do not present any information of the structure and monomer sequence in the material.30 Thioacidolysis offers an estimate of lignin monomer units derived only from 8−O−4 structures.31 Since extractability and digestibility are affected not only by the amount of lignin but also by its monomer composition and the way the monomers are arranged in the plant cell wall, it is important to accurately determine this information when trying to develop less recalcitrant biomass. Much information on the structure of lignin has been obtained via artificial lignin, synthesized in vitro by polymerization of lignin precursors using oxidizing agents.32 Therefore, we have pursued this line of reasoning by establishing a simple way to polymerize lignin in vitro using different proportions of the three monomers and commercial peroxidases to mimic the natural polymerization process. The resulting compounds can be analyzed via ultrahigh performance liquid chromatography (UPLC) coupled to tandem mass spectrometry (MS/MS) to determine their structure and build a library of lignin oligomer data. Then, soluble lignin found in plant tissues can be gently extracted without degradation and its sequence and structure determined by UPLC−MS/MS in comparison to this library. The monomers and small oligomers that compose the soluble lignin found in plant tissues will later be incorporated into cell wall lignin. Information on the proportions of S, G, and H monomers, as well as the types of linkages found in soluble lignin, should be proportionally represented in cell wall lignin. This information may be correlated to the cell wall composition and provide an unbiased profile of the monomers and types of linkages found in the plant sample. Other phenolic compounds could also be incorporated into the in vitro lignin structure, and
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EXPERIMENTAL SECTION Oligomer Synthesis and Extraction. The procedure for the in vitro synthesis of oligomers was adapted from de Angelis et al.33 In brief, 5 mg of individual monomers [p-coumaryl alcohol (H), coniferyl alcohol (G), sinapyl alcohol (S), SigmaAldrich, U.S.A.] or mixtures (3 mg of each monomer) were dissolved in 2 mL of sodium phosphate buffer pH 6.5 (10 mmol·mL−1) containing 27 mmol·L−1 of (CTA)2SO4. Then, 3% hydrogen peroxide (67 μL) and 3 enzyme units of horseradish peroxidase types I or II (Sigma, U.S.A.) were added. The reaction was allowed to proceed at 30 °C for 5 min or 1 or 2 h and then interrupted with a few drops of a 5% solution of Na2S2O7. Monomers and oligomers were extracted with ethyl acetate, which was recovered, washed with a saturated aqueous NaCl solution, and finally evaporated under a stream of N2. The dry residue was dissolved in an aqueous solution containing 35% acetonitrile prior to analysis. Ultrahigh Performance Liquid Chromatography− Tandem Mass Spectrometry (UPLC−MS/MS). Each sample (5 μL) was injected into an Acquity UPLC coupled to a TQD triple-quadrupole mass spectrometer (Micromass-Waters, Manchester, England). The chromatographic separation was carried out using a Waters Acquity C18-BEH (2.1 mm × 50 mm, 1.7 μm) column and gradient elution ranging from 5% to 100% acetonitrile (solvent B) in 8 min, and using Milli-Q water with 0.1% formic acid as solvent A. A flow of 0.200 mL·min−1 and column temperature of 30 °C were used. Electrospray ionization in the negative ion mode was used under the following conditions: capillary 3.0 kV and cone 50 V, ion source temperature 150 °C, desolvation temperature 300 °C. The MS/ MS spectra were obtained by collision-induced dissociation (CID) with collision energy of 20 V. During the development of the chromatographic method, ammonium acetate buffer, ammonium hydroxide, and formic acid were tested, as well as ionization in both positive and negative ion modes. Due to the acid character of the phenolic monomers and oligomers, negative ion mode ionization was the most suitable. The resulting spectra were analyzed to determine the structure of the oligomers and, when possible, compared to data found in literature Extraction and Analysis of Soluble Lignin from Sugar Cane. Approximately 0.1 g of freeze-dried and finely ground sugar cane sample was extracted with 1 mL of a solution containing 80% ethanol in water in an ultrasonic bath for 30 min and then centrifuged in a benchtop centrifuge at full speed to eliminate debris. The supernatant was collected and evaporated in a speed vac. The dry material was resuspended in 0.5 mL of Milli-Q water, and lignin was extracted with 0.6 mL of ethyl acetate. The organic phase was collected and evaporated under a stream of N2. The dry residue was solubilized in an aqueous solution containing 35% acetonitrile prior to analysis. The deprotonated ions of the lignin oligomers 7016
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chromatograms and MS/MS of all these compounds may be found in the Supporting Information (Figure S1). The difference of 30 Da between the H, G, and S monomers (due to the addition of an OCH3 group in positions 3 and 5) allows us to determine which units come from which monomer. Another important point to note is that β-aryl ether links (8− O−4) result in an increment of 18 Da in the mass of the lignin molecule, because a molecule of water is incorporated as an (8−O−4) link is formed between monomers. The mechanism of this reaction is discussed in detail by Morreel et al.30 Further lignin chain elongation by (8−O−4) links results in the incorporation of one more water molecule per link. Several oligomers with β-aryl ether (8−O−4), phenylcoumaran (8−5), and resinol (8−8) links (Figure 2) were characterized based on their characteristic fragmentation patterns in comparison to literature.30 Monomers. Deprotonated coniferyl aldehyde (m/z 177) fragments by a loss of a methyl radical (15 Da) due to the presence of a methyl group and a further loss of CO (28 Da). Deprotonated sinapyl aldehyde (m/z 207) fragments by two subsequent losses of methyl radicals (15 Da each) followed by a loss of CO (28 Da). The deprotonated p-coumaryl, coniferyl, and sinapyl alcohols (m/z 149, 179, and 209, respectively) lose H2O (18 Da), and the latter also lose one and two methyl radicals, respectively, due to the methoxy groups in positions 3 and 5 (Figure 1). These fragmentation patterns were observed for authentic standards of these compounds as well as for the compounds identified in the sugar cane sample. Dimers. The fragmentation pattern of the deprotonated G− G dimers, G(8−5)G and G(8−8)G (m/z 357), and G(8−O− 4)G (m/z 375), were compared to data from literature.30 Fragmentation of the deprotonated (8−5) linked dimer yields a fragment ion of m/z 221 which is characteristic of the 4aliphatic end. For the deprotonated (8−8) linked dimer, the fragment ion of m/z 151 is characteristic of the 8-phenolic end. As the mass of the (8−O−4) linked dimers increases by 18 Da due to the incorporation of a molecule of water, their deprotonated molecules present characteristic fragment ions of m/z 179 for the 4-aliphatic end and m/z 195 for the 8phenolic end. Two deprotonated H−H dimers, H(8−5)H and H(8−8)H (m/z 297), were detected. For the deprotonated H(8−5)H dimer a characteristic loss of water (18 Da) and a fragment ion of m/z 191 for the 4-aliphatic end (similar to the m/z 221 ion of the G(8−5)G dimer) are observed, as well as a fragment ion of m/z 121 for the 8-phenolic end. The difference of 30 Da between H and G monomers is due to the lack of methoxy groups on the H monomer. The only other dimer found incorporating H was G(8−5)H; its deprotonated molecule (m/ z 327) forms a characteristic fragment ion (m/z 191), indicating the H is on the 4-aliphatic end. Two deprotonated S−G dimers, S(8−5)G and S(8−8)G (m/z 387), were observed. The difference of 30 Da between S and G monomers is due to the presence of one more methoxy group on the S monomer, but the fragmentation pattern of these S−G dimers parallels that of the respective G−G dimers. Fragmentation of the deprotonated (8−5) linked dimer yields a fragment ion of m/z 221 which is characteristic of G on the 4aliphatic end. The S(8−O−4)G dimer is 18 Da heavier than other S−G dimers, and its deprotonated molecule (m/z 405) yields a fragment ion of m/z 225 indicating that the S unit is on the 8-phenolic end. The inverse structure, G(8−O−4)S, was not detected.
were identified based on the comparison of their retention time, m/z, and MS/MS with data from our DIY library.
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RESULTS AND DISCUSSION Building the Library. Two different peroxidase enzymes and three different reaction times were tested to determine which would produce the longest chains and greatest variety of oligomers. The results in Table 1 show that both enzymes and Table 1. Retention Time and m/z of the Deprotonated Molecules of Monolignols, Aldehydes, and Synthesized Oligomers, as well as the Type of Peroxidase and Reaction Time Useda m/z
retention time (min)
structure
peroxidase type
2.70 3.3 2.72 3.26 2.75 3.7 3.91 3.21 3.39
p-coumaryl alcohol (H) coniferyl aldehyde coniferyl alcohol (G) sinapyl aldehyde sinapyl alcohol (S) H(8−5)H H(8−8)H G(8−5)H G(8−5)G
1 1 1 1
4.08
G(8−8)G
1 and 2
375
3.76
G(8−O−4)G
1 and 2
387
3.42
S(8−5)G
1 and 2
3.9
S(8−8)G
1 and 2
405 417
3.2 3.79
S(8−O−4)G S(8−8)S
1 and 2 1 and 2
435 493 535 553 571 583
3.91 3.74 4.58 3.7 3.54 3.57
S(8−O−4)G H(8−5)H(8−O−4)G G(8−5)G(8−5)G G(8−O−4)G(8−5)G G(8−O−4)G(8−O−4)G G(8−O−4)S(8−5)G
1 2 1 1 1 1
601 613
3.66 3.58 3.52, 4.34
S(8−O−4)G(8−5)G G(8−O−4)G(8−O−4)S S(8−O−4)S(8−5)G
1 and 2 1 and 2 1 and 2
3.87, 4.34
G(8−O−4)S(8−8)S
1 and 2
3.65 3.82 3.24 4.77
S(8−O−4)G(8−O−4)S S(8−O−4)S(8−8)S S(8−O−4)S(8−O−4)S S(8−O*−4)S(8−8)S(8−O− 4)Sb
1 1 1 1
149 177 179 207 209 297 327 357
631 643 661 851
and and and and
2 2 2 2
and 2 and and and and
and and and and
2 2 2 2
2 2 2 2
reaction time
1 and 2 h 1 and 2 h 1 and 2 h 5 min and 1 h 5 min and 1 h 5 min and 1 h 5 min and 1 h 5 min and 1 h 1h 5 min and 1 h 1h 1h 1h 1h 1h 5 min and 1 h 1 and 2 h 1h 5 min and 1 h 5 min and 1 h 1h 1h 2h 1h
a
Only deprotonated molecules were observed in the negative ion mode. bAsterisk denotes an unusual type of 8−O−4 coupling.
quite variable reaction times often produced similar oligomers, demonstrating this synthesis to be robust. Twenty-five compounds were identified: most were dimers and trimers, and one tetramer. The retention times of the monomers, and their aldehydes, were also determined. Small amounts of coniferyl aldehyde and sinapyl aldehyde were found as contaminants of the respective monomer standards. The 7017
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Figure 2. Dimers of G units showing the three main links. Their characteristic fragmentation patterns are indicated by red lines. (I) The 8−O−4, βaryl ether link yields fragment ions A− (m/z 195) and B− (m/z 179). (II) The 8−5, phenylcoumaran link yields fragment ion B− (m/z 221). (III) The 8−8, resinol link yields fragment ion A− (m/z 151).
Figure 3. General structures of the trimers identified by UPLC−MS. The (8−O−4) link present in structures I, II, and III results in fragment ions A− and BC−. A− has the mass of the deprotonated monomer plus a molecule of water, and the BC− fragment ion has the same m/z as the dimers composed of S and G (Table 1). Structure IV can only have two 8−5 links as there is no increase in the mass of the monomers, and fragment ion C− is typical of the phenylcoumaran link.
fragment ions: A− the 8-phenolic end presents the mass of the deprotonated monomer plus a molecule of water, and BC− the 4-aliphatic end, which corresponds to the remaining deprotonated dimer (Figure 3). The MS/MS spectrum of the trimers also yields fragment ions which are characteristic of the monomers which make up the remaining dimer, allowing the elucidation of the complete structure. A deprotonated trimer composed of two H and one G unit was observed, H(8−5)H(8−O−4)G (m/z 493). The fragment ion of m/z 179 indicates that the 4-aliphatic end (B) is the G monomer and the 8-phenolic end (A) is composed of two H units. The fragment ion of m/z 191 is typical of a cleaved phenylcoumaran (8−5) link indicating that the two H units are linked this way.
Two S−S dimers were observed. Deprotonated S(8−8)S (m/z 417) can only have the 8−8 link because of the methoxy group blocking the 5-position. The S(8−O−4)S dimer is 18 Da heavier than the S(8−8)S dimer due to water incorporation, and its deprotonated molecule (m/z 435) yields the fragment ions of m/z 225 and 209, which are characteristic of this structure. Trimers. The deprotonated molecules of the following trimers were detected and identified by comparison to data from literature:30 G(8−O−4)G(8−O−4)G (m/z 571), G(8− O−4)S(8−5)G (m/z 583), S(8−O−4)G(8−5)G (m/z 583), S(8−O−4)S(8−5)G (m/z 613), G(8−O−4)S(8−8)S (m/z 613), and S(8−O−4)S(8−8)S (m/z 643). Deprotonated trimers which begin with the (8−O−4) link are easily recognized as the C8−O link is cleaved, forming two distinct 7018
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from our library. The structures found in the sugar cane sample are listed in Table 2.
Two G−G−G trimers, not described in literature, were also detected. In the deprotonated G(8−O−4)G(8−5)G trimer (m/z 553) the fragment ion of m/z 195 indicates that the 8phenolic end (A) is a G, and the fragment ion of m/z 357 indicates that the (B) group is composed of two G units. Furthermore the fragment ion of m/z 339 (B − H2O) indicates that these units have a phenylcoumaran (8−5) link. The other deprotonated trimer, G(8−5)G(8−5)G (m/z 535), yields the following fragment ions: m/z 177 and m/z 221 of the 4aliphatic end indicate that only 8−5 coupling is present. The deprotonated G−G−S trimer, G(8−O−4)G(8−O−4)S (m/z 601), only has (8−O−4) links. The fragment ion of m/z 195 indicates that the 8-phenolic end (A) is a G, and the fragment ion of m/z 209 shows that the remaining dimer is G(8−O−4)S. Another deprotonated trimer, S(8−O−4)G(8− O−4)S (m/z 631), also has exclusively (8−O−4) links. Fragment ion m/z 225 indicates that the 8-phenolic end (A) is an S unit, and m/z 209 indicates that the 4-aliphatic end is also an S. The deprotonated S−S−S trimer, S(8−O−4)S(8−O−4)S (m/z 661), has, as expected, only 8−O−4 links, because the other positions are blocked by methoxy groups. The fragment ion of m/z 225 confirms that the 8-phenolic end (A) is an S, and fragment ion of m/z 209 confirms that the 4-aliphatic end is also an S. Tetramer. Only one tetramer, formed exclusively by S units, was found: S(8−O−4)S(8−O*−4)S(8−8)S. The fragmentation pattern of this deprotonated oligomer suggests that one of the (8−O−4) links is atypical (highlighted by an asterisk), similar to the one previously described by de Angelis et al.33 The fragment ion of m/z 417 indicates that there is an (8−8) link, and the fragment ion of m/z 225 is characteristic of an (8− O−4) link on the 8-phenolic end. The fragment ion of m/z 207 indicates that the atypical (8−O*−4) link is in the middle of the oligomer. Therefore, we propose the structure presented in Figure 4.
Table 2. Structures Identified in the Soluble Lignin Extracted from Sugarcane Listed by the m/z of the Deprotonated Molecules m/z
structure
177 179 207 209 327 357 375 387 405 417 583 613 643
coniferyl aldehyde coniferyl alcohol (G) sinapyl aldehyde sinapyl alcohol (S) G(8−5)H G(8−5)G and G(8−8)G G(8−O−4)G S(8−5)G and S(8−8)G S(8−O−4)G S(8−8)S G(8−O−4)G(8−5)G S(8−O−4)S(8−5)G S(8−O−4)S(8−8)S
Although a larger number of samples and a more complete study are necessary to characterize lignin composition in sugar cane, this proof-of-principle analysis demonstrates that it is perfectly feasible to build your own lignin database and use it to study plant samples. In the sugar cane sample we found only one structure containing the H monomer. This might be expected since in monocotyledons similar amounts of S and G units at found, but much lower amounts of H units are reported.9 However, depending on the lignin source, the DIY library needs further enrichment by incorporation of new structures identified in plant samples, but basic structures and fragmentation patterns can be easily obtained by the approach presented here. Furthermore, both easily cleaved (8−O−4) links as well as more chemically recalcitrant links are observed, showing that all these types of chemical links are already present in the oligomers before being incorporated in the cell wall. Once lignin is incorporated in cell walls, aggressive treatments are needed to extract it, which necessarily result in chemical modification of this complex biopolymer. Therefore, there are several methods used to evaluate lignin composition, but no consensus. By evaluating the easily extracted soluble lignin, valuable information regarding overall lignin composition (and therefore the recalcitrance of plant biomass before further processing) may be gleaned.
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Figure 4. Structure of the tetramer identified by UPLC−MS. The m/z of the deprotonated molecule is compatible with four S units plus one water molecule. Fragment ions of m/z 225 (A−) and 625 (BCD−) indicate that one (8−O−4) unit is on the 8-phenolic end. The fragment ion of m/z 417 indicates that the other extremity has an S(8−8)S structure (CD−). For this reason we propose that B is linked to C (green box) via an atypical 8−O−4 link, because no water molecule was incorporated. Upon fragmentation, however, the CD− structure, typical of β-aryl ether links, is formed.
CONCLUSION A robust method to build a DIY lignin oligomer library was tested. This library can be built using commercially available enzymes, standards, and reagents, and it is relatively easy to accomplish. In the present study only the main monomers, pcoumaryl alcohol (H), coniferyl alcohol (G), and sinapyl alcohol (S), were tested, but other phenylpropanoids can be incorporated in the synthetic lignin in the same manner. Soluble lignin was easily extracted from the plant material with an ethanolic solution and then transferred to ethyl acetate by liquid/liquid extraction. This method is extremely soft in comparison to the aggressive treatments routinely used to extract lignin from cell walls and does not alter the structure of the oligomers like those treatments.
Soluble Lignin from Sugar Cane. Extracted soluble lignin from sugar cane internodes was used to validate the DIY library. The retention time of the chromatographic peaks, as well as the m/z of the deprotonated lignin molecules and monomers and their tandem mass spectra (MS/MS), were compared with data 7019
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(16) Boyce, C. K.; Zwieniecki, M. A.; Cody, G. D.; Jacobsen, C.; Wirick, S.; Knoll, A. H.; Holbrook, N. M. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 17555−17558. (17) Atanassova, R.; Favet, N.; Martz, F.; Chabbert, B.; Tollier, M.T.; Monties, B.; Fritig, B.; Legrand, M. Plant J. 1995, 8, 465−477. (18) Bernard-Vailhé, M.-A.; Cornu, A.; Robert, D.; Maillot, M.-P.; Besle, J.-M. J. Agric. Food Chem. 1996, 44, 1164−1169. (19) Sewalt, V. J. H.; Ni, W.; Jung, H. G.; Dixon, R. A. J. Agric. Food Chem. 1997, 45, 1977−1983. (20) Goujon, T.; Ferret, V.; Mila, I.; Pollet, B.; Ruel, K.; Burlat, V.; Joseleau, J.-P.; Barrière, Y.; Lapierre, C.; Jouanin, L. Planta 2003, 217, 218−228. (21) Ralph, J.; Akiyama, T.; Kim, H.; Lu, F.; Schatz, P. F.; Marita, J. M.; Ralph, S. A.; Reddy, M. S. S.; Chen, F.; Dixon, R. J. Biol. Chem. 2006, 281, 8843−8853. (22) Jackson, L. A.; Shadle, G. L.; Zhou, R.; Nakashima, J.; Chen, F.; Dixon, R. A. BioEnergy Res. 2008, 1, 180−192. (23) Li, X.; Weng, J.-K.; Chapple, C. Plant J. 2008, 54, 569−581. (24) Fu, C.; Mielenz, J. R.; Xiao, X.; Ge, Y.; Hamilton, C. Y.; Rodriguez, M.; Chen, F.; Foston, M.; Ragauskas, A.; Bouton, J.; Dixon, R. A.; Wang, Z.-Y. Proc. Natl. Acad. Sci. U.S.A. 2011, 108, 3803−3808. (25) Grabber, J. H.; Hatfield, R. D.; Lu, F.; Ralph, J. Biomacromolecules 2008, 9, 2510−2516. (26) Harris, D.; DeBolt, S. Plant Biotechnol. J. 2010, 8, 244−262. (27) Campbell, M. M.; Sederoff, R. R. Plant Physiol. 1996, 110, 3−13. (28) Lapierre, C.; Monties, B.; Rolando, C.; Laboratoire de Chirale. J. Wood Chem. Technol. 1985, 5, 277−292. (29) Brinkmann, K.; Blaschke, L.; Polle, A. J. Chem. Ecol. 2002, 28, 2483−2501. (30) Morreel, K.; Dima, O.; Kim, H.; Lu, F.; Niculaes, C.; Vanholme, R.; Dauwe, R.; Goeminne, G.; Inzé, D.; Messens, E.; Ralph, J.; Boerjan, W. Plant Physiol. 2010, 153, 1464−1478. (31) Kishimoto, T.; Chiba, W.; Saito, K.; Fukushima, K.; Uraki, Y.; Ubukata, M. J. Agric. Food Chem. 2010, 58, 895−901. (32) Sarkanen, K.; Ludwig, C. Lignins: Occurrence, Formation, Structure and Reactions; Sarkanen, K. V., Ludwig, C. H., Eds.; WileyInterscience, 1971. (33) De Angelis, F.; Nicoletti, R.; Spreti, N.; Verì, F. Angew. Chem., Int. Ed. 1999, 38, 1283−1285.
A UPLC−MS/MS method for the separation and characterization of monomers and oligomers was developed and was equally applicable to the synthetic lignin and to the soluble lignin extracted from a sample of sugar cane. The oligomers found in the sugar cane sample were indicative of the monomers available (mainly G and S) and the types of links between them, which directly influence the recalcitrance of plant material to further processing. Therefore, the composition of soluble lignin could be used to determine the overall composition and recalcitrance of biomass. The UPLC−MS/MS method was used qualitatively in the present study. Due to the structural similarity of the monomers, calibration curves using one of the monomers as a standard could give a reasonably accurate quantification of other oligomers, based on the comparison of their chromatographic peak areas. Further studies will evaluate a larger number of sugar cane samples and characterize the lignin composition of diverse varieties.
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ASSOCIATED CONTENT
S Supporting Information *
Additional information as noted in text. This material is available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was made possible through FAPESP BIOEN Grant 2008/58035-6. A.C.H.F.S. and P.M. thank CAPES and CNPq for research fellowships, respectively.
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REFERENCES
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dx.doi.org/10.1021/ac301112y | Anal. Chem. 2012, 84, 7015−7020
